Significance

RNA-protein granules play crucial roles in cell biology, development, and disease. Yet their molecular assembly and biochemical functions remain poorly understood. This work focuses on a granule scaffold protein, called PGL, to gain molecular insights into granule assembly and function. We identify a dimerization domain in a PGL region required for granule assembly and determine its crystal structure. The structure reveals a positively charged channel of the right size for binding single-stranded RNA. However, we instead find that PGL is a base-specific RNA endonuclease. The importance of this work lies in its molecular clarification of a building block for granule assembly and discovery of an activity that provides a new view of a scaffold protein as an enzyme.

Abstract

Cellular RNA-protein (RNP) granules are ubiquitous and have fundamental roles in biology and RNA metabolism, but the molecular basis of their structure, assembly, and function is poorly understood. Using nematode “P-granules” as a paradigm, we focus on the PGL granule scaffold protein to gain molecular insights into RNP granule structure and assembly. We first identify a PGL dimerization domain (DD) and determine its crystal structure. PGL-1 DD has a novel 13 α-helix fold that creates a positively charged channel as a homodimer. We investigate its capacity to bind RNA and discover unexpectedly that PGL-1 DD is a guanosine-specific, single-stranded endonuclease. Discovery of the PGL homodimer, together with previous results, suggests a model in which the PGL DD dimer forms a fundamental building block for P-granule assembly. Discovery of the PGL RNase activity expands the role of RNP granule assembly proteins to include enzymatic activity in addition to their job as structural scaffolds.

Cytoplasmic RNA-protein (RNP) granules are found in virtually all cells and are thought to be central to RNA metabolism (1, 2). These diverse organelles include P-bodies, stress granules, neuronal granules, and germ granules (2). RNP granules are not membrane-bound and display liquid–liquid phase-separation properties (3, 4). Many of their molecular components have been identified, including scaffold proteins: proteins that recruit other key granule components and are sufficient to induce RNP granule assembly. Major challenges now are to understand how RNP granules are assembled and how they control RNAs.

Germ granules are exemplary RNP granules with a profound yet largely mysterious role in metazoan germ-line development. These granules possess common components across phyla (5) but use unique scaffold proteins, such as Drosophila Oskar (6), zebrafish Bucky Ball (7), and Caenorhabditis elegans paralogs PGL-1 and PGL-3 (8, 9), called PGL collectively. Germ granule scaffold proteins from different phyla have distinct amino acid sequences with no conserved domains. The importance of these scaffolds has been attributed to their function in germ granule assembly (for examples, see refs. 10⇓–12). However, the molecular basis of that assembly and how it impacts RNA regulation remain unknown.

Here we focus on the Caenorhabditid PGL scaffold proteins and their role in assembly of nematode germ granules, called P-granules (13). P-granules are required for germ-line survival (8, 9) and germ-line totipotency (14). A recent model proposes that P-granules capture selected mRNAs exiting the nucleus (15), an idea based on the finding that untranslated mRNAs are enriched in P-granules, but translated mRNAs are absent (15, 16). The consequences of that capture are unclear but may include mRNA repression.

The PGL family comprises the closely related PGL-1 and PGL-3 proteins plus divergent PGL-2. PGL-1 and PGL-3 are required for adult germ cell development but the function of PGL-2 is unknown (8). All three PGL proteins interact with each other in vitro (8), but thus far only PGL-1 and PGL-3 are known to self-assemble into granules when expressed in nematode somatic cells or in mammalian cell culture (17, 18). By primary sequence prediction, PGLs have only one recognizable region, C-terminal RGG repeats (Fig. 1A) (8, 9), which are associated with protein turnover and RNA binding (17, 19). In non-PGL proteins, RGG repeats can recruit RNA binding proteins (20) and facilitate in vitro granule formation (21). Experiments in tissue culture cells implicate the N-terminal half of the PGL protein—but not the RGG repeats—in P-granule assembly (17). Whereas full-length PGL-3 self-assembled, a mutant lacking ∼160 residues from the PGL-3 N-terminal half (Fig. 1A) no longer formed granules (17). Thus, in-roads have been made but molecular detail about how PGL organizes itself into a granule is lacking.

Here we identify a PGL dimerization domain (DD) and propose that PGL dimers are a key building block for P-granule assembly. We determine the PGL-1 DD crystal structure and find a novel 13-helix fold that creates a positively charged channel as a homodimer. Testing the idea that this channel might bind RNA, we discovered that PGL DD is an RNase and determined PGL-1 DD’s specificity in vitro for guanosine in single-stranded RNAs. We suggest that PGL DD has a dual role in P-granules, as a domain essential for assembly and as an RNase.

Results

PGL Dimerization and Its Crystal Structure.

We biochemically characterized the PGL proteins to better understand regions contributing to granule assembly. Amino acid sequence alignments identified a large region spanning the N-terminal two-thirds of the protein that was conserved among Caenorhabditids (Fig. 1A and Materials and Methods). Our initial characterization focused on purified recombinant C. elegans PGL-3 (Ce-PGL-3) (Fig. S1A). Ce-PGL-3 residues 1–447 ran in the void on a sizing column (Fig. 1B) and multimerized in chemical cross-linking experiments (Fig. S1B), consistent with that fragment assembling into a large multimer. Limited proteolysis of Ce-PGL-3 residues 1–447 identified a single protected fragment (Fig. S1C). N-terminal sequencing and additional proteolysis mapped the fragment to residues 205–447 (Fig. 1A). This 25-kDa domain dimerized in solution, as determined by both size-exclusion chromatography (Fig. 1B) and chemical cross-linking (Fig. S1D). We therefore refer to this region as the PGL DD.

To gain molecular insight into PGL DD dimerization, we determined crystal structures of C. elegans (Ce) and Caenorhabditis remanei (Cr) PGL-1 DD to 3.6 Å and 1.6 Å, respectively (Table S1; more details in Materials and Methods). Both structures revealed a domain of 13 α-helices that assume an identical, novel fold (Fig. 2A and Fig. S2 A and B) (RMSD = 0.826 Å). A similar structure could not be found with protein fold-alignment software (Dali) (22), consistent with primary sequence alignments reporting PGLs as novel proteins. Cr- and Ce-PGL-1 DD both crystallized as single subunits in their asymmetric unit (Fig. 2A and Fig. S2 A and B). Therefore, the asymmetric unit on its own could not be used to identify the biologically relevant dimer. We instead analyzed the crystal packing of both structures to see whether the dimer crystallized on a crystal symmetry axis. Our two crystal structures are of different PGL-1 homologs and have completely different space groups, but have only one conserved protein–protein interface between them (Fig. 2B and Fig. S2C). A similar dimerization-fold was calculated by PISA (23), an assembly-prediction program. Thus, both proteins crystallized as identical homodimers on different twofold crystal symmetry axes, providing convincing evidence that this is the authentic dimer structure.

The PGL-1 DD dimer makes extensive subunit contacts. For example, the higher resolution Cr-PGL-1 DD dimer interface has a large buried surface area (1,239 Å2) that is predicted to form a combined total of 13 hydrogen bonds and salt bridges (PISA) (Fig. 2C). Importantly, the dimer-interface residues are highly conserved between PGL-1 and PGL-3 homologs (Fig. 2D). However, those residues are not as well conserved in PGL-2 (Fig. S2D).

PGL-1 DD dimerization forms a central channel between its subunits (Fig. 2B and Fig. S2C). The N-terminal half of PGL-1 DD makes most of the dimer interactions (10 of 13 total hydrogen bonds and salt bridges, Cr-PGL-1 DD) to form the top of the channel. In contrast, helix α11 in the C terminus makes minimal contacts with its counterpart in the adjacent subunit (3 of 13 total hydrogen bonds and salt bridges, Cr-PGL-1 DD) to enclose the channel at the bottom (Fig. 2B and Fig. S2C). The asymmetry between the number of contacts at the top and bottom of the channel implies that the relatively weak α11:α11 interaction may be dynamic and permit access to the channel interior without disrupting dimerization. The channel diameter, measured from C-α traces of its surrounding helices, is roughly 15 Å, providing sufficient room for single-stranded—but not double-stranded—nucleic acid. Moreover, the electrostatic surface potential of the channel is basic (Fig. 2E). We postulate that the PGL-1 DD dimer may accommodate RNA within its channel, given the channel size, surface charge, and the established relationship between P-granules and RNA. Thus, the PGL-1 DD structure provides insight into its dimerization and also suggests a possible second role, previously not considered, as an RNA binding domain.

PGL-1 Is a Guanosine-Specific Endonuclease.

To test the idea that PGL DD binds RNA, we used EMSA, in which stable RNA–protein complexes migrate more slowly than free RNA. However, the opposite was observed. When Ce-PGL-1 DD was incubated with 5′ labeled RNAs, the RNA migrated faster than RNA alone (Fig. 3A). The faster RNA migration on native gels implied a decrease in RNA size, which was validated on denaturing gels (Fig. 3C). Thus, PGL-1 DD cleaves RNA. RNA cleavage was also found for Ce-PGL-3 and Cr-PGL-1 (Fig. S3 A and B), demonstrating conservation of this enzymatic activity among closely related PGL proteins.

Ce-PGL-1 DD cleaved certain RNAs but not others (Fig. 3 A–D), implying specificity for sequence, or secondary or tertiary structure. Incubation of Ce-PGL-1 DD with a longer and more complex RNA (pos-1 3′UTR, 315 bases) yielded a cleavage pattern similar to that of RNase T1 (Fig. 3E), a guanosine-specific RNA endonuclease that cleaves 3′ to guanosines (24). PGL-1 DD incubation with a polyuridine RNA oligo harboring a single interior guanosine base (polyU/G) enriched for a single cleavage product identical in size to that produced by RNase T1 (Fig. 3F). In contrast, incubation with RNase A, a pyrimidine-specific RNase (25), caused complete degradation (Fig. 3F). Inclusion of commercial RNase inhibitors suppressed enzymatic activity of RNase T1 and A, but had no observable effect on PGL DD cleavage (Fig. 3F), arguing against an RNase contaminant being responsible for the PGL DD cleavage result. These findings, along with our crystal structures showing a novel fold, suggest that PGL DD is a new endonuclease.

The size similarity between the cleavage products of PGL-1 DD and T1 endonucleases led us to explore further the PGL-1 DD RNase specificity. We tested the ability of PGL-1 DD to cleave a polyuridine RNA harboring other single-base changes within the oligo. No RNA cleavage was observed upon inclusion of uridine, cytidine, and adenosine bases (Fig. 3G), whereas cleavage occurred in a concentration-dependent manner with guanosine (Fig. 3G and Fig. S3C). Because the uridine base pairs with adenosine, we also tested a polyadenine RNA oligo with an interior cytosine base (polyA/C) but similarly observed no cleavage (Fig. S3D). We next explored ways to inhibit PGL-1 DD cleavage of its guanine-containing substrate. RNA endonucleases cleave RNA using its 2′ hydroxyl, not found in DNA, for nucleophilic attack (26). PGL-1 DD could not cleave a DNA polyU/G oligo, nor an RNA polyU/G oligo with a 2′ hydroxyl modified to 2′ fluorinated guanosine (Fig. 3H). Therefore, PGL-1 DD specifically cleaves guanosine-containing RNA. PGL DD may have additional, unexplored RNA target-sequence specificity other than guanosine, and may be like RNase T1, whose catalytic rates are greatly affected by the base adjacent to guanosines (27). PGL binding partners, particularly those that bind RNA, such as IFE-1 (12), may modify the PGL DD RNase activity. We next tested for secondary-structure specificity. The diameter of the PGL DD dimer channel is predicted to fit only single-stranded RNA (see above), assuming no significant structural rearrangement. In support of a restriction to single-stranded RNA, cleavage was also blocked by preincubation of polyU/G with its complementary oligo (polyA/C) to form double-stranded RNA (Fig. 3I). Addition of a noncomplementary polyuridine RNA oligo had no effect on cleavage (Fig. 3I). Taken together, our results suggest that PGL-1 DD is a guanosine-specific, single-stranded RNA endonuclease.

What residues mediate the PGL DD enzymatic activity? Ribonucleases have a diversity of domains that form active sites with a similar molecular composition (26). Cleavage is typically accomplished through acid/base chemistry via amino acid side chains or divalent metals (26). Cleavage by PGL-1 DD is not likely to rely on a common divalent metal, because all cleavage assays were done in the presence of a metal chelator (EDTA) and addition of common metals (Mg, Mn) did not significantly affect cleavage rates (Fig. S4A). Other metals (e.g., Co, Cu) caused protein precipitation and could not be adequately tested. In metal-independent RNases, like RNase T1, histidine typically serves as the base in the active site, and is paired with either another histidine or an acidic side chain (26). We found only a single conserved histidine paired with a neighboring acidic side chain, a glutamate (Fig. S4 B–D). Mutation of this histidine reduced RNA cleavage activity significantly (Fig. S4E), but mutation of its pairing glutamate had no effect (Fig. S4E). Therefore, PGL DD is unlikely to have a classic RNase active site.

We sought other residues that might mediate catalysis. At the top of the channel, within a solvent-exposed region of the dimer subunit interface (Fig. 4 A and B), a conserved glutamine in one subunit comes together with its counterpart in the other subunit (Fig. 4 A–C). Mutation of that glutamine to alanine (Ce-PGL-1 Q342A) abolished PGL-1 DD cleavage activity (Fig. 4D) but did not affect dimerization (Fig. S5). Moreover, unlike the wild-type protein, incubation of the PGL-1 Q342A mutant protein with polyU/G slowed migration of the labeled oligo (Fig. 4E). This slower migrating band likely represents a PGL-1 DD–RNA complex and indicates that PGL-1 Q342A binds RNA. We conclude that PGL-1 Q342A abrogates RNase activity without affecting dimerization or RNA binding. The glutamine could be part of the active site, coordinate an untested metal, or affect function allosterically through local misfolding. Access to Q342 requires either dimer subunit separation or entry into the channel. We speculate therefore that the dimer channel is crucial for RNA interaction and enzyme catalysis. Regardless, the PGL-1 Q342A mutant demonstrates that PGL-1 DD is responsible for the observed RNase activity, and excludes the notion that this activity is a contaminant from recombinant protein purification or the environment.

Purification of MBP-tagged PGL-1 DD. (A) Wild-type and mutant PGL-1 DD both dimerize. Sizing column of recombinant MBP::Ce-PGL-1 DD wild-type (WT) and Q342A mutant protein. Column void volume (“void”) and free MBP (“MBP”) labeled with arrows. Black bar indicates fractions pooled and used in RNA cleavage analyses. (B) Coomassie-stained SDS/PAGE gel of purified MBP::Ce-PGL-1 wild-type and mutant proteins E442Q, H478A, and Q342A. Protein pooled from sizing column fractions (A, black bar). The wild-type samples represent two different protein preparations (WT 1, WT 2). (C) MBP::Ce-PGL-1 DD wild-type (WT) and Q342A recombinant protein purified from a sizing column rerun on the same column one day later. No protein is observed in the void, and both WT and Q342A elute at a similar position to that observed in the initial sizing column run. A minor peak is again observed later during the elution and attributed to MBP.

To test the role of PGL-1 RNase activity in vivo, we used CRISPR/Cas9 gene editing to generate two independent but identical Q342A RNase defective mutations in the endogenous C. elegans pgl-1 locus (Materials and Methods). Wild-type worms were fertile at 20 °C and 26.5 °C; in contrast pgl-1–null mutants were fertile only at 20 °C but sterile at 26.5 °C (Fig. 4F) (9). Unlike the pgl-1–null mutant, both pgl-1 Q342A mutants were fertile at both 20 °C and 26.5 °C (Fig. 4F). Therefore, PGL-1 RNase activity is likely not required for its role in fertility.

Discussion

This work reveals two key features of PGL proteins: the PGL dimerization domain and PGL-1 guanosine-specific RNA endonuclease activity. Discovery of the PGL dimerization domain allows us to expand on the Hanazawa model for P-granule assembly (17) to include PGL DD as a fundamental building block of the P-granule scaffold. Hanazawa et al. found a PGL deletion that eliminates granule assembly (Fig. 1A) (17), and we now know that their deletion removes part of PGL DD. Putting PGL dimerization (present work) together with PGL multimerization (17), we now suggest that the P-granule framework is constructed from multimers of PGL dimers. This strategy is similar in nature to that of Oskar, the fly germ granule scaffold protein that also dimerizes to facilitate assembly (28).

Discovery of the PGL RNase activity changes our view of RNP granule scaffold proteins. Previously, only the RGG repeats linked the PGL scaffold with RNA (17). The PGL DD RNA endonuclease activity was unexpected. Its structure assumes a novel fold and lacks any cluster of amino acids recognizable as a classic RNase active site, which opens a host of questions about its enzyme mechanism, base specificity, P-granule function, regulation, and conservation. PGL’s enzymatic activity is modest compared with classic guanosine-specific RNases, like RNase T1. PGL could have additional sequence specificity not yet identified, or specificity for a modified guanosine, like the 5′ cap. Alternatively, inefficiency may be ideal for a granule-forming enzyme to permit RNase activity only when present at high concentrations within P-granules.

Enzymatic activities in other germ granule scaffold proteins, zebrafish Bucky Ball and Drosophila Oskar, have not been identified, but it is plausible that they, like PGL, may contain novel enzymatic domains or recruit enzymes to serve analogous roles. Intriguingly, Maelstrom, a Piwi-interacting RNA (piRNA) biogenesis factor and germ-granule component, was recently identified as a novel guanosine RNase (29). The RNA targets of the Maelstrom nuclease are unknown and its enzymatic activity is dispensable in vivo for piRNA biogenesis. Although Maelstrom and PGL are structurally unrelated, their parallels are striking: both reside in germ granules, both possess guanosine RNA endonuclease activity, and that enzymatic activity is dispensable in vivo. An attractive idea is that convergent evolution established RNase activity in distinct proteins within germ granules and that their activities serve a common purpose in reproduction.

What might the PGL RNase do in P-granules? Selected mRNAs localize to P-granules and that localization correlates with their translational inhibition (16). PGL DD RNase may cleave the 5′ cap or 3′ regulatory regions of mRNAs retained in P-granules, and hence block translation. Potential targets include genes associated with neuronal and muscle cell development, which are inhibited in P-granules to prevent aberrant germ cell differentiation (14). The RNases responsible for piRNA biogenesis are largely unknown and those that are known have no obvious homologs (30). For example, Zucchini, a piRNA biogenesis RNase, has no known nematode equivalent. PGL may be the functional counterpart of Zucchini or play some other role in piRNA metabolism. Regardless, the insights from this work provide a starting point to further explore the molecular assembly and RNA regulatory mechanisms of this model RNP granule.

Protein Purification.

Details regarding protein purification can be found in SI Materials and Methods. Purification was finished on a S200 size-exclusion column (GE Healthcare) in PGL buffer [20 mM Hepes pH 7.0, 50 mM NaCl, 0.5 mM Tris(2-carboxyethyl)phosphine pH 7.0 (TCEP, Sigma)]. FPLC fractions were again analyzed by SDS/PAGE and Coomassie staining. Peak fractions were concentrated with a 10K cut-off Amicon Ultra-4 concentrator (EMD Millipore) and stored at 4 °C until use. Final protein concentration was estimated by A280. See Fig. S5B for an example of the final protein used in RNA experiments.

Protease digestion.

Proteomics-grade Trypsin (1 μg/mL final concentration; Sigma-Aldrich) was added to recombinant PGL-3 residues 1–447 (0.4 mg/mL) and samples were taken at 5, 10, 20, 40, and 60 min, and before the addition of trypsin as a negative control. Samples were analyzed by SDS/PAGE and Coomassie staining. SDS/PAGE gels were also transferred to PVDF, and submitted for N-terminal sequencing (Tufts University Core Facility, M. Berne).

pos-1 and RNA oligo digestion.

Purified 32P-labeled pos-1 3’ UTR RNA was incubated with C. elegans MBP::PGL-1 DD (1 and 3 μM), RNase T1 (1.2 nM; Thermo Scientific), RNase A (0.02 nM; Sigma-Aldrich), and a negative control (dH20) in cleavage buffer for 1 h at room temperature (∼20 °C). Samples were phenol-chloroform extracted, and run on a 0.75-mm 15% (vol/vol) urea-TBE gel (National Diagnostics). Gels were fixed, dried, and developed similar to the native gels. 32P-labeled PolyU/G RNA oligonucleotide (Table S2) was incubated with C. elegans MBP::PGL-1 DD (1 μM), RNase T1 (1.2 nM), RNase A (0.02 nM), and a negative control (dH20) in cleavage buffer for 1 h at room temperature. Samples were phenol-chloroform–extracted and run on a 20% (vol/vol) urea-TBE gel and analyzed similar to that described in pos-1 RNA digestion analysis. Both experiments were repeated at least three times with similar results.

Time-course digestion assays.

32P-labeled RNA and DNA oligonucleotides (1 nM, based on oligo concentration before 32P-labeling) were incubated with varying concentrations of C. elegans MBP::PGL-1 DD (3, 1, 0.3, 0.1 μM) in cleavage buffer with RNasin RNase Plus Inhibitor (1 Unit/μL; Promega). 32P-labeled PolyU/G RNA oligonucleotide was also incubated with 3 μM C. elegans MBP::PGL-1 DD mutants (E442Q, H478A, Q342A). Samples were taken at increasing time points (5, 10, 15, 20, 30, 45, 60, 90, 120 min) and reaction stopped with phenol-chloroform. A no-enzyme sample (dH20) was incubated 120 min in parallel. A “0” time point represents a sample taken after addition of labeled RNA or DNA to the reaction. Samples were phenol-chloroform extracted and run in sample buffer on 15% (vol/vol) TBE-Urea 0.75-mm mini gels. Gels were fixed, dried, and imaged similar to the pos-1 and RNA oligo digestion assays. Band intensities were measured using ImageQuant and data presented as (“cleavage product”)/(“full length oligo” + “cleavage product”). Graphs were generated by Excel and represent an average of three separate experiments.

Double-stranded RNA cleavage assay.

32P-labeled PolyU/G RNA oligonucleotide (1 nM, based on oligo concentration before 32P-labeling) was incubated with unlabeled PolyA/C and PolyU RNA oligonucleotides (Table S2) at 1:10, 1:1, 1:0.1, and 1:0 molar ratios based upon the concentration of PolyU/G used in the initial labeling reaction. Reactions were performed, and samples processed and analyzed similar to time-course assays.

Protease digestion.

Proteomics-grade trypsin (1 μg/mL final concentration; Sigma-Aldrich) was added to recombinant PGL-3 residues 1–447 (0.4 mg/mL) and incubated for 60 min at room temperature (∼20 °C). Samples were taken at 5, 10, 20, 40, and 60 min, and before the addition of trypsin as a negative control. Samples were analyzed by SDS/PAGE and Coomassie staining. SDS/PAGE gels were also transferred to PVDF, and the 25-kDa cleavage product observed submitted for N-terminal sequencing (Tufts University Core Facility, M. Berne). The C terminus of the cleavage product was estimated to be the end of the recombinant protein. Trypsin incubation with the recombinant PGL-3 DD residues 205–447 did not create observable cleavage product.

Protein Crystallization and Structure Determination.

Recombinant proteins were initially screened on 96-well sitting-drop trays set by the Mosquito (TTP Labtech) at 20 °C. C. elegans and C. remanei PGL-1 DD and PGL-3 DD all had crystal “hits” identified, but only the C. elegans and C. remanei PGL-1 crystals could be reproduced in 24-well, 4-μL (2 μL protein:2 μL mother liquor) hanging-drop trays. The initial C. elegans PGL-1 DD crystals were small and numerous. This condition was further tested with an Additive Screen (Hampton Research). Several additives improved crystal size while decreasing nucleation. The largest, most reproducible crystals were observed with the addition of guanidine HCl (GuCl). Final crystal conditions for C. elegans PGL-1 DD were 1.575–1.625 M sodium malonate pH 5.9, 50–100 mM GuCl, 1 mM TCEP, 1 mM sodium azide. At their largest, the hexagonal-shaped crystals were ∼20 μm × 20 μm × 80 μm. Crystals were observed at 2 wk and grew in size until ∼4 wk. Crystals were frozen in the described condition with 20% (vol/vol) ethylene glycol. The dataset submitted was from a crystal soaked in gadolinium chloride (10 mM) overnight before freezing. Diffraction data were collected at LS-CAT and GM/CA. Spots were observed to 2.7 Å, but a complete dataset could only be collected to 3.6 Å in space group P61 2 2.

Data processing was performed in HKL2000 (33), model building done in Coot (34), and refinement in PHENIX (35). Phase extension of the Thimerosal-soaked (mercury, Hg) and native datasets could not be achieved, presumably because of the change in cell dimensions. Instead, we phased the Hg dataset (HySS) (40, 41) and Phaser-EP (36), built an initial C. remanei model (∼50% of the final structure) from the Thimerosal-soaked dataset, and used this model to obtain phases in the native dataset. The final C. remanei model was used for model phases in the C. elegans dataset. Molecular replacement was performed by Phaser (36). In the C. remanei native dataset map, water molecules were identified by PHENIX, ArpWarp (42), and individual placement and refinement. Unaccounted for densities, too large to be water, were later identified. Placement of sulfate and ethylene glycol molecules in these densities improved Rwork and Rfree. Final structure statistics are found in Table S1. Datasets, structure models, and additional information can be found in the RCSB Protein Data Bank. The PDB IDs are as follows: C. remanei PGL-1 DD, native (PBD ID code 5COW); C. remanei PGL-1 DD, Thimerosal (Hg) (PBD ID code 5CV3); C. elegans PGL-1 DD (PBD ID code 5CV1).

Nucleic Acid Digest Assays.

Nucleic acid labeling.

RNA and DNA oligos were commercially synthesized (Table S2) (IDT Technologies). The 5′ labeling with 32P was done with phosphonucleotide kinase (New England Biolabs) and EasyTide 32P γ-ATP (Perkin-Elmer). Briefly, 5 pmol oligo was labeled for 1 h under the manufacturer’s conditions, and purified from unincorporated 32P with an illustra MicroSpin G-25 column (GE Healthcare). 32P labeled oligo was diluted further based upon initial labeling concentrations. A plasmid of pos-1 mRNA with 3′UTR was cloned from mixed-stage N2 nematode TRIzol-extracted total RNA with the SuperScript III First-Strand Synthesis Kit (Life Technologies). The PCR product was cloned into a vector using the Zero Blunt TOPO PCR Cloning Kit (Life Technologies). The 3′UTR (315 bases) (Table S2) was PCR-amplified by Phusion Polymerase with primers that included a T7 promoter (AATACGACTCACTATAGGGAGATTTCTCTCGTCGAAATTTCTGATC; TGCTGATTACGAGAAATTTCATTTTATG). The PCR product was treated with DpnI (New England Biolabs), gel-purified (Qiagen), and transcribed using the AmpliScribe T7-Flash Transcription Kit (Epicentre) with GTP in rate-limiting quantities to promote incorporation of 32P α GTP (Perkin-Elmer). After 2 h, the resulting product was separated from unincorporated nucleotides using NucAway Spin columns (Life Technologies) and run on an 8% (vol/vol) TBE-Urea polyacrylamide gel (UreaGel System, National Diagnostics). The RNA transcript band was detected by autoradiography, excised, gel-purified, phenol-chloroform–extracted, and ethanol-precipitated. The final product was reconstituted in dH20 for use.

Time-course digestion assays.

32P-labeled RNA and DNA oligonucleotides (1 nM, based on oligo concentration before 32P-labeling) were incubated with varying concentrations of C. elegans MBP::PGL-1 DD (3, 1, 0.3, 0.1 μM) in cleavage buffer with RNasin RNase Plus Inhibitor (1 Unit/μL; Promega). 32P-labeled PolyU/G RNA oligonucleotide was also incubated with 3 μM C. elegans MBP::PGL-1 DD mutants (E442Q, H478A, Q342A). Samples were taken at increasing time points (5, 10, 15, 20, 30, 45, 60, 90, 120 min) and reaction stopped with phenol-chloroform. A no-enzyme sample (dH20) was incubated 120 min in parallel; we occasionally observed a smaller cleavage product that we attributed to contaminating RNase digestion. A “0” time point represents a sample taken after addition of labeled RNA or DNA to the reaction. Samples were phenol-chloroform–extracted and run in sample buffer on 15% (vol/vol) TBE-Urea 0.75-mm mini gels. Gels were fixed, dried, and imaged similar to the pos-1 and RNA oligo digestion assays. Band intensities were measured using ImageQuant and data presented as (“cleavage product”)/(“full length oligo” + “cleavage product”). Graphs were generated by Excel and represent an average of three separate experiments.

Double-stranded RNA cleavage assay.

32P-labeled PolyU/G RNA oligonucleotide (1 nM, based on oligo concentration before 32P-labeling) was incubated with unlabeled PolyA/C and PolyU RNA oligonucleotides (Table S2) at 1:10, 1:1, 1:0.1, and 1:0 molar ratios based upon the concentration of PolyU/G used in the initial labeling reaction. Mixtures were heated for 5 min at 70 °C and cooled at room temperature (∼20 °C) for 10 min before the addition of 1 μM C. elegans MBP::PGL-1 DD, or a no enzyme control, in cleavage buffer. Reactions were incubated for 30 min at room temperature and stopped with phenol-chloroform. Samples were processed and imaged on 15% (vol/vol) TBE-Urea gels, and analyzed similar to time-course assays. Graphs were generated by Excel and represent three separate experiments.

CRISPR/Cas9 genome editing.

The CRISPR/Cas9 coconversion genome editing approach (38) was used to generate a Q342A mutation in endogenous pgl-1. Briefly, an sgRNA construct containing the U6 promoter and sgRNA scaffold from pDD162 (43) along with the targeting sequence gtcagagcggaagtctttcc was cloned into the XmaI site of pUC19 using one step isothermal DNA assembly (32) to generate the clone pJK1889. The repair template was a 980-nt single-strand DNA oligo (tcctctctgggattcctacgagtgtcagagcggaggttttcccaggactggccagaagtgtctacaagagtgcggtgttcctcggcaatcacatcatc) that inserted the Q342A mutation as well as an MscI restriction site. Injections were carried out using young wild-type hermaphrodite C. elegans with DNA concentrations, as described previously (38), and F1 rollers were screened for the desired mutation by PCR and MscI digest. Two alleles, q842 and q843, were recovered from separate injected animals, and therefore represent independent editing events. Mutants were verified by Sanger sequencing.

Fertility assays.

Wild-type or homozygous pgl-1 mutant (bn102, q842, q843) late L4/young adults were placed singly on fresh plates at 20 °C and allowed to generate embryos for 1 d. These parents were then transferred singly to fresh plates, shifted to 26.5 °C, and allowed to generate embryos for an additional 1 d, after which time they were killed. Their progeny were then scored as young adults by dissecting scope, as described previously (8). Briefly, if animals contained at least one healthy looking embryo on either of 2 d, they were scored as fertile and then immediately killed. If they did not contain at least one healthy looking embryo on either of 2 d, they were scored as sterile.

Acknowledgments

The authors thank M. Preston and C. Valley for training; M. Cox for equipment; A. Helsley-Marchbanks for help preparing the manuscript; L. Vanderploeg for help with the figures; and members of the J.K. laboratory, K. Desai, E. Montemayor, T. Nguyen, R. Raines, A. Hoskins, S. Butcher, D. Updike, and S. Strome for helpful discussions. Use of the Life Sciences-Collaborative Access Team Sector 21 was supported by the Michigan Economic Development Corporation and the Michigan Technology Tri-Corridor (Grant 085P1000817). The National Institute of General Medical Sciences and National Cancer Institute Structural Biology Facility at the Advanced Photon Source has been funded in whole or in part with Federal funds from the National Cancer Institute (ACB-12002) and the National Institute of General Medical Sciences (AGM-12006). This research used resources of the Advanced Photon Source, a US DOE Office of Science User Facility operated for the DOE Office of Science by the Argonne National Laboratory under Contract DE-AC02-06CH11357. The Berkeley Center for Structural Biology is supported in part by the National Institutes of Health (NIH), National Institute of General Medical Sciences, and the Howard Hughes Medical Institute. The Advanced Light Source is supported by the Director, Office of Science, Office of Basic Energy Sciences, of the US DOE under Contract DE-AC02-05CH11231. S.T.A. was supported by the Eunice Kennedy Shriver National Institute of Child Health & Human Development of the NIH under Awards F32HD071692 and K99HD081208; C.A.B. was supported by NIH Grants GM094584, GM094622, and GM098248; and M.W. was supported by NIH Grant GM50942. J.K. is an Investigator of the Howard Hughes Medical Institute.

Blood-sucking sand flies from disparate global regions have a predilection for feeding on the marijuana plant (Cannabis sativa), and the findings hint at a potential avenue for controlling sand flies, which can transmit leishmaniasis.